Fluorescent energy transfer probes are an important tool in genetic analysis. These probes, also known as dual-labeled probes (DLPs) or self-quenching probes, are generally comprised of a fluorescent donor (a fluorophore) and a quencher linked to an oligonucleotide. This basic design, wherein a signal change is detected once the probe hybridizes to its intended target, is used in a variety of biological applications.
One method for detecting hybridization using fluorophores and quenchers is to link fluorescent donors and quenchers to a single oligonucleotide such that there is a detectable difference in fluorescence when the oligonucleotide is unhybridized as compared to when it is hybridized to its complementary sequence. In so-called molecular beacons, a partially self-complementary oligonucleotide is designed to form a hairpin and is labeled with a fluorescent donor at one end of the molecule and a quencher at the other end (U.S. Pat. No. 5,925,517). Intramolecular annealing to form the hairpin brings the donor and quencher into close proximity for fluorescent quenching to occur. Intermolecular annealing of such an oligonucleotide to a target sequence disrupts the hairpin, which increases the distance between the donor and quencher and results in a detectable increase in the fluorescent signal of the donor.
Oligonucleotides are not required to form a hairpin structure for this method to work efficiently. The fluorophore and quencher can be placed on an oligonucleotide such that when it is unhybridized and in a random coil conformation, the quencher is able to quench fluorescence from the fluorophore (U.S. Pat. No. 5,538,848). Once the oligonucleotide hybridizes to a complementary nucleotide sequence it becomes more extended and the distance between the fluorophore and quencher is increased, resulting in reduced quenching and increased fluorescence.
Oligonucleotides labeled in a similar manner can also be used to monitor the kinetics of PCR amplification. In one version of this method, commonly known as a 5′-nuclease cleavage or hydrolysis assay, an oligonucleotide probe is designed to hybridize to the target sequence on the 3′ side (“downstream”) of one of the amplification primers. During PCR, the 5′-3′ exonuclease activity of the DNA polymerase digests the 5′ end of the probe thereby separating the fluorophore from the quencher. The fluorescence intensity of the sample increases as an increasing number of probe molecules are digested during the course of amplification (U.S. Pat. No. 5,210,015).
DLPs find use in other molecular/cellular biology and diagnostic assays, such as in end-point PCR, in situ hybridizations, in vivo DNA and RNA species detection, single nucleotide polymorphism (SNPs) analysis, enzyme assays, and in vivo and in vitro whole cell assays (see Dirks and Tanke, Biotechniques 2006, 40:489-486; Bustin, Journal of Molecular Endocrinology 2002, 29:23-39; Mackay, Clin Microbiol Infect, 2004, 10:190-212).
In one mechanism of fluorescence quenching termed ground state quenching, the fluorophore and the quencher associate to form a ground state complex which is not fluorescent. For ground state quenching to occur there need not be spectral overlap between the fluorophore and the quencher.
The most common mechanism of fluorescent quenching is fluorescence resonance energy transfer (FRET). In FRET, energy transfer occurs through space by dipolar coupling between the fluorophore and quencher and requires that there be overlap between the emission spectrum of the fluorescent donor and the absorbance spectrum of the quencher. This requirement complicates the design of probes that utilize FRET because quenchers are limited in their effective wavelength range. For example, the quencher known as BHQ-1, which absorbs light in the wavelength range of about 500-550 nm, quenches fluorescent light emitted by fluorescein, which fluoresces maximally at about 520 nm, but is of limited utility for Texas Red (emission maximum=615) or Cy5 (emission maximum=670). In contrast, the quencher BHQ-3, which absorbs light in the wavelength range of about 650-700 nm is almost completely ineffective at quenching fluorescein but is effective at quenching Cy5. In general, the number of quenchers that are known to be capable of quenching the fluorescence of any given fluorophore is limited.
Although fluorescent dyes themselves can be employed to quench fluorescence from other dyes, preferred quenchers will not fluoresce (or minimally fluoresce) so that background fluorescence is minimized. These quenchers are commonly referred to as dark quenchers. Dark quenchers allow for an increased signal to noise ratio in assays that employ DLPs, resulting in increased sensitivity. In addition, the lack of secondary fluorescence facilitates the use of additional fluorophores in multiplexed assay formats which utilize multiple distinct probes each containing a different fluorophore. If a quencher emitted light in a certain region, then additional probes could not bear fluorophores that emit light in that same region.
A number of factors are considered in designing a self-quenching probe. These include the ease of synthesis, the compatibility of the fluorophore and quencher, duplex stability, and the specificity of the probe in hybridizing to the intended target.
Duplex stability between complementary nucleic acid molecules is frequently expressed as the “melting temperature”, Tm, of the duplex. Roughly speaking, the Tm indicates the temperature at which a duplex nucleic acid dissociates into two single strands. Nucleic acid hybridization is generally performed at a temperature slightly below the Tm, so that hybridization between a probe or primer and its target nucleic acid is optimized, while minimizing non-specific hybridization of the probe or primer to other, non-target nucleic acids. Duplex stability and Tm are also important in applications, such as PCR, where thermocycling may be involved. During such thermocycling melting steps, it is important that the sample temperature be raised sufficiently above the Tm so that duplexes of the target nucleic acid and its complement are dissociated. In subsequent steps of reannealing, however, the temperature must be brought sufficiently below the Tm that duplexes of the target nucleic acid and primer are able to form, while still remaining high enough to avoid non-specific hybridization events. For a general discussion, see Rychlik et al., Nucleic Acids Research 1990, 18:6409-6412.
Shorter oligonucleotides can help increase the specificity of a primer or probe, allowing for the discrimination of even a single mismatch between the probe and a potential complementary target. The shorter the oligonucleotide, the greater the effect of a single-base mismatch on duplex stability. However, the disadvantage of using such short oligonucleotides is that they hybridize weakly, even to a perfectly complementary sequence, and thus must be used at lower temperatures, which are unfavorable for reactions that use thermal stable enzymes, such as PCR. Certain modified nucleosides such as locked nucleic acids (LNAs) (U.S. Pat. No. 7,060,809) and C5-propynyl pyrimidines (U.S. Pat. No. 5,484,908) can be incorporated into oligonucleotides to increase duplex stability. Many nucleoside analogs, however, especially those having bulkier substituents attached to the base, are destabilizing. For example, fluorescein-dT can destabilize a duplex by up to 4° C. (Bioorganic & Medicinal Chemistry Letters, 13:2785-2788 2003).
Modified nucleosides employed to increase Tm are typically placed internally within an oligonucleotide sequence replacing a natural base. In contrast, non-nucleoside substituents when introduced internally within an oligonucleotide, either as a replacement for a base or as an insertion between bases, generally interfere with hybridization. For example, insertion of an abasic fluorescein group into an oligonucleotide has been observed to destabilize a duplex by 2-4° C. (DNA Seq. 4:135-141, 1993).
There are several classes of compounds that are known to increase binding affinity between complementary nucleic acid strands. One class is major groove binders, which includes proteins or ligands that bind to the major groove (the wider groove around a DNA helix). A second class, minor groove binders (MGBs), include non-covalently bound and covalently bound compounds. Because the minor groove of a helix is narrower in A-T rich regions, some noncovalently bound MBGs recognize the shape of the helix and preferably bind to specific sequence regions. For example, netropsin and distamycin preferably bind to A-T regions (see Bailly and Henichart, ACS, vol. 2, 379-393 (1991). Covalently bound MGBs (U.S. Pat. No. 6,084,102) are typically linked to the 5′ or 3′ end of oligonucleotides (U.S. Pat. App. 2009/0259030) and are known to increase binding affinity and allow for shorter length probes.
A third class, intercalators, are generally flat polycyclic compounds, examples being acridine or lipticine derivatives (see U.S. Pat. No. 4,835,263). Intercalating compounds stabilize a duplex by fitting in between the bases of the nucleic acid monomers. They can be covalently or noncovalently bound. Some minor groove compounds, such as 4′,6-diamidino-2-phenylindole (DAPI), also intercalate.
Another group of compounds, capping reagents, are terminally attached compounds that favor Watson-Crick duplexes by stacking on the terminal base pair (Dogan, Z. et al., J. Am. Chem. Soc. 2004, 126, 4762-4763). Such groups include stilbene derivatives (Wu, T. et al., J. Am. Chem. Soc. 1995, 117, 8785-8792) and pyrenylmethylpyrrolindol (Narayanan, S. et al., Nucleic Acids Res. 2004, 32, 2901-2911).
The efficiency of quenching through FRET is extremely sensitive to the distance between the fluorophore and quencher (RFQ), varying with the reciprocal of RFQ to the sixth power. Maximally efficient quenching minimizes background fluorescence and improves the sensitivity of the 5′-nuclease assay and other hybridization assays in which DLPs are used. Generally, for ease of synthesis and to avoid disruption of hybridization of the probe to the target sequence, the dye and quencher are attached to the ends of the oligonucleotide. For the 5′-nuclease assay, the most common configuration is to attach the dye at the 5′-end of the oligonucleotide and the quencher at the 3′-end. DLPs used in the 5′-nuclease assay are typically 25 to 30 bases in length. Even with the use of Tm enhancing modifications, such as LNA bases or a minor groove binder, probe length is still usually 14 to 18 bases. Any method that permits placement of the fluorophore and quencher in closer proximity within a probe without destabilizing the duplex formed between the probe and its target nucleic acid will improve quencher efficiency and enhance the performance of the probe.